Invasive Species Compendium

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Datasheet

Mycosphaerella pini
(Dothistroma blight)

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Datasheet

Mycosphaerella pini (Dothistroma blight)

Summary

  • Last modified
  • 03 October 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Natural Enemy
  • Preferred Scientific Name
  • Mycosphaerella pini
  • Preferred Common Name
  • Dothistroma blight
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Fungi
  •     Phylum: Ascomycota
  •       Subphylum: Pezizomycotina
  •         Class: Dothideomycetes
  • Summary of Invasiveness
  • Dothistroma septospora (the anamorphic form of the fungus) has spread rapidly around the world since its identification as a serious crop pathogen in Tanzania in 1957, and is now globally widespread. The fungus is believed to be endemic to pines in C...

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Pictures

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PictureTitleCaptionCopyright
Mycosphaerella pini (Dothistroma blight); attached needles of Pinus nigra infected by M. pini (Dothistroma septospora).
TitleSymptoms
CaptionMycosphaerella pini (Dothistroma blight); attached needles of Pinus nigra infected by M. pini (Dothistroma septospora).
Copyright©Leo Pehl
Mycosphaerella pini (Dothistroma blight); attached needles of Pinus nigra infected by M. pini (Dothistroma septospora).
SymptomsMycosphaerella pini (Dothistroma blight); attached needles of Pinus nigra infected by M. pini (Dothistroma septospora).©Leo Pehl
Mycosphaerella pini (Dothistroma blight); typical 'red bands' on needles of Pinus nigra.
TitleSymptoms
CaptionMycosphaerella pini (Dothistroma blight); typical 'red bands' on needles of Pinus nigra.
Copyright©Leo Pehl
Mycosphaerella pini (Dothistroma blight); typical 'red bands' on needles of Pinus nigra.
SymptomsMycosphaerella pini (Dothistroma blight); typical 'red bands' on needles of Pinus nigra.©Leo Pehl
Mycosphaerella pini (Dothistroma blight); fructifications, rupturing needle epidermis of Pinus mugo.
TitleFructifications
CaptionMycosphaerella pini (Dothistroma blight); fructifications, rupturing needle epidermis of Pinus mugo.
Copyright©Leo Pehl
Mycosphaerella pini (Dothistroma blight); fructifications, rupturing needle epidermis of Pinus mugo.
FructificationsMycosphaerella pini (Dothistroma blight); fructifications, rupturing needle epidermis of Pinus mugo.©Leo Pehl
Cross-section through a conidioma of Dothistroma septospora, staining: Thionine.
TitleConidioma
CaptionCross-section through a conidioma of Dothistroma septospora, staining: Thionine.
Copyright©Leo Pehl
Cross-section through a conidioma of Dothistroma septospora, staining: Thionine.
ConidiomaCross-section through a conidioma of Dothistroma septospora, staining: Thionine.©Leo Pehl
Mycosphaerella pini (Dothistroma blight); conidia of Dothistroma septospora.
TitleConidia
CaptionMycosphaerella pini (Dothistroma blight); conidia of Dothistroma septospora.
Copyright©Leo Pehl
Mycosphaerella pini (Dothistroma blight); conidia of Dothistroma septospora.
ConidiaMycosphaerella pini (Dothistroma blight); conidia of Dothistroma septospora.©Leo Pehl
Mycosphaerella pini (Dothistroma blight); cross-section through an ascoma. Staining: Thionine
TitleAscoma
CaptionMycosphaerella pini (Dothistroma blight); cross-section through an ascoma. Staining: Thionine
Copyright©Leo Pehl
Mycosphaerella pini (Dothistroma blight); cross-section through an ascoma. Staining: Thionine
AscomaMycosphaerella pini (Dothistroma blight); cross-section through an ascoma. Staining: Thionine©Leo Pehl
Mycosphaerella pini (Dothistroma blight); asci and ascospores. Staining: Cotton blue in lactic acid.
TitleAsci and ascospores
CaptionMycosphaerella pini (Dothistroma blight); asci and ascospores. Staining: Cotton blue in lactic acid.
Copyright©Leo Pehl
Mycosphaerella pini (Dothistroma blight); asci and ascospores. Staining: Cotton blue in lactic acid.
Asci and ascosporesMycosphaerella pini (Dothistroma blight); asci and ascospores. Staining: Cotton blue in lactic acid.©Leo Pehl

Identity

Top of page

Preferred Scientific Name

  • Mycosphaerella pini Rostr. 1957

Preferred Common Name

  • Dothistroma blight

Other Scientific Names

  • Cytosporina septospora Dorog. 1911
  • Dothistroma pini Hulbary 1941
  • Dothistroma septosporum (Dorog.) Morelet 1968
  • Eruptio pini (Rostr.) M.E. Barr 1996
  • Scirrhia pini A. Funk & A.K. Parker 1966
  • Septoriella septospora (Dorog.) Sacc. 1931

International Common Names

  • English: blight: pine; brown needle blight: pine; circular: persimmon leaf spot; Dothistroma needle blight; needle blight: pine; pine blight; pine brown needle blight; pine needle blight; pine red band needle blight; red band needle blight:pine; red-band disease; red-band fungus; red-band needle blight
  • Spanish: estriado roio de las aciculas del pino; manchas circulares del caqui; manchas circulares del palosanto
  • French: maladie des bandes rouges du pin; strie rouge des aiguilles du pin; taches circulaires du kaki

Local Common Names

  • Germany: Dothistroma-Nadelbraeune: Kiefer; Nadelbraeune: Kiefer

EPPO code

  • SCIRPI (Mycosphaerella pini)

Summary of Invasiveness

Top of page Dothistroma septospora (the anamorphic form of the fungus) has spread rapidly around the world since its identification as a serious crop pathogen in Tanzania in 1957, and is now globally widespread. The fungus is believed to be endemic to pines in Central America and Nepal (Evans, 1984; Ivory, 1994), where it is a foliar pathogen. The pathogen is particularly damaging where trees are planted out of their host range, most notably in the Southern hemisphere where large commercial monocultures of susceptible species such as Pinus radiata have been planted in New Zealand and Chile. At the same time the disease has increased in importance in the USA, where it has caused damage to shelterbelt, amenity and Christmas tree crops of P. nigra, P. ponderosa and P. contorta. Plant disease reports from Europe also suggest an increase in the prevalence in that part of the world. The pathogen has probably spread by a combination of factors: transport of infected planting material, and wind/cloud dissemination of spores between land masses (Gibson, 1974). The sexual form of the fungus, Mycosphaerella pini, does not have such a wide host range and is largely restricted to the Northern hemisphere. Moreover, since the asexual rather than the sexual spores are thought to be the primary source of inoculum, it is likely that the teleomorph is less invasive than the anamorph.

Taxonomic Tree

Top of page
  • Domain: Eukaryota
  •     Kingdom: Fungi
  •         Phylum: Ascomycota
  •             Subphylum: Pezizomycotina
  •                 Class: Dothideomycetes
  •                     Subclass: Dothideomycetidae
  •                         Order: Capnodiales
  •                             Family: Mycosphaerellaceae
  •                                 Genus: Mycosphaerella
  •                                     Species: Mycosphaerella pini

Notes on Taxonomy and Nomenclature

Top of page The anamorphic form of M. pini was first described in Russia as Cytosporina septospora (Doroguin, 1911). Hulbary (1941) named the fungus responsible for an outbreak of needle blight in Illinois, USA, as Dothistroma pini. Morelet (1968) considered these fungi to be identical and made a new combination Dothistroma septospora (Dorog.) Morelet, a nomenclature accepted by Sutton (1980). However, the synonym Dothistroma pini is still in common use.

Three varieties of the conidial state are recognized on the basis of conidial length. Thyr and Shaw (1964) distinguished Dothistroma pini var. pini with conidial lengths of 15.4-28.0 (mean 22.4) µm and Dothistroma pini var. linearis with conidial lengths of 23.0-42.0 (31.9) µm. The long-spored variety linearis is reported to occur in western States of the USA and Canada whilst the short-spored variety pini is found in the central and eastern States of North America, and in England, New Zealand, Australia and Chile (Ivory, 1967; Peterson and Graham, 1974; Edwards and Walker, 1978). A third variety with intermediate conidial lengths of 13.0-47.5 (28.9) µm occurring in Africa (predominantly Kenya) was named by Ivory (1967) as Dothistroma pini var. keniensis. Sutton (1980) lists the conidial lengths of these varieties under their respective synonyms as: D. septospora var. septospora 12.5-32.5 (22) µm; D. septospora var. lineare 20.0-67.5 (37.5) µm; D. septospora var. keniense 15-47.5 (29) µm. The distinctness of these varietal divisions has been questioned (Funk and Parker, 1966; Sutton, 1980). Gadgil (1967) found large variations in conidial length and shed doubt on the value of conidial length as a diagnostic character, suggesting that separate varieties should not be recognized. Ivory (1967) found that conidia from isolates in culture were generally larger than those collected from diseased needles and noted inconsistencies in measurements of conidial lengths from the same sample depending on whether or not they were incubated in a damp chamber for 5 days. Similarly, more recent studies have shown no clear distinction between these varieties on the basis of conidial length or of internal transcribed sequence (ITS) DNA sequence analysis (Edwards and Walker, 1978; Roux, 1984; Bradshaw et al., 2000). Evans (1984), who carried out a thorough and comprehensive study on a global collection of isolates, does not support the division into varieties.

The genus Mycosphaerella is considered by some to be polyphyletic, as more than 40 anamorph genera are associated with it (Goodwin et al., 2001). Barr (1996) separated species with Dothistroma and Lecanosticta anamorphs into a new genus, Eruptio, on the assumptions that these two anamorphs are closely related and are different from other species within Mycosphaerella. However, phylogenetic analysis of internal transcribed sequence (ITS) data contradicted these assumptions and suggested that the genus Mycosphaerella is monophyletic. The teleomorph name for Dothistroma septospora should remain within Mycosphaerella (Goodwin et al., 2001).

Description

Top of page In diseased needles the most visible part of the fungus is usually the asexual fruiting bodies. In some countries sexual fruiting bodies can also be seen. Colour illustrations of diseased needles, asexual and sexual stages of M. pini can be found in Pehl and Wulf (2001).

Fruiting bodies are initially white and subepidermal but becoming brown-black, acervular and erumpent as they develop, eventually breaking through the host epidermis and cuticle and leaving torn flaps around the fruiting bodies. Conidiomata are circular to oval, 300-1500 µm long, 300 µm wide. However, the conidiomata are of varying complexity and this appears to be controlled by host and climate (Evans, 1984). Sometimes referred to as a pycnidium, the asexual fruiting structure is an elongate subepidermal acervulus produced within a well-defined stroma. However in some needle samples (particularly from resistant hosts) only shallow open stromatic acervuli are found.

A dense layer of hyaline, elongated conidiophores (20-40 x 2-2.5 µm) develop on the upper surface. Conidia are formed from the apex of the conidiophores and on short side branches. In shallow acervuli the fertile area is restricted to a short palisade of conidiogenous cells and integrated conidiophores are not seen as they rapidly become pseudoparenchymatic. Moreover in older erumpent acervuli conidiophores are converted into stromatic tissue until only the apical cells remain (Edwards and Walker, 1978; Evans, 1984).

Conidia are exuded in a white or pale pink mucilaginous mass, are hyaline or very faintly tinted, filiform, straight or slightly curved, (1-) 3 (-5) septate, and 15-36 x 2-3 µm, although wider variations in spore sizes are known.

Sutton (1980) lists the conidiospore lengths of the three varieties as follows:

D. septospora var. septospora, syn. Dothistroma pini var. pini, conidia 12.5-32.5 (mean 22) x 2.5-4 (3) µm
D. septospora var. lineare, syn. Dothistroma pini var. linearis, conidia 20-67.5 (37.5) x 2-3 (2.5) µm
D. septospora var. keniense, syn. Dothistroma pini var. keniensis, conidia 15-47.5 (29) x 1.5-4 (2.5) µm

Ascomata are typically aggregated in red bands on diseased pine needles, subepidermal, becoming erumpent, black, uniloculate to multiloculate, up to 850 µm wide and composed of dark-brown pseudoparenchyma. The ascogonia with trichogynes and the spermagonia are produced in separate stromata. Trichogynes usually form in a small stroma beneath the host tissue surface whilst spermatia usually form in erumpent stromata in which macroconidia are often also formed. Trichogynes are brown, septate, 36-100 µm long and 4-5 µm wide; ascogonia are brown, coiled or flexuous and approximately 20 µm long and up to 6 µm wide. Spermatiferous cells are found in columnar chains in locules; spermatia (microconidia) are rod-shaped, hyaline, 1.5-2.5 x 0.5-1 µm and embedded in mucous (Funk, 1979). Asci are saccate to cylindrical, bitunicate, 35-55 x 5-9 µm, 8-spored, hyaline. Ascopores are elliptic, 1-septate, 8-16 x 3-4 µm, hyaline.

In culture, M. pini is slow-growing (colony diameter of 8-15 mm after 28 days at 21°C on malt extract agar) and exudes a reddish-brown pigment (dothistromin) into the agar. Hyphae of aerial mycelium are hyaline to olive-brown, whilst hyphae of the substrate mycelium are darker brown. Macroconidia and microconidia can be formed in culture, generally in a white or pinkish conidial slime. The overall morphologies of the colonies in culture, their growth rates and the amounts of dothistromin exuded into the agar, are highly variable between isolates (Bradshaw et al., 2000).

Distribution

Top of page The anamorph Dothistroma septospora has a global distribution, with most widespread occurrence in countries that grow susceptible pine species out of their native habitat on a commercial scale (e.g. Chile, New Zealand). The teleomorph has a more limited distribution and is predominantly found in the Northern hemisphere, including Canada (Funk and Parker, 1966), the USA (Cobb and Miller, 1968; Peterson and Graham, 1974; Peterson and Harvey, 1976), Germany (Butin and Richter, 1983), Yugoslavia (Karadzic, 1989), Poland (Kowalski and Jankowiak, 1998) and Portugal (Fonseca and Laflamme, 1997). Intriguingly, in countries where M. pini has been a major needle pathogen (e.g., Chile, East Africa, Australia, New Zealand) the teleomorph has not been reported (Evans, 1984).

M. pini disease is commonly found at high-altitude sites, such as native P. mugo at 1200-1600 m (Maschning and Pehl, 1994) and P. radiata at 2500-3000 m (Evans and Oleas, 1983). In a survey of Central America, Evans did not find the disease below 1500 m (Evans, 1984).

See also CABI/EPPO (1998, No. 219).

Distribution Table

Top of page

The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

BhutanPresentCABI/EPPO, 2010; EPPO, 2014
Brunei DarussalamPresentCABI/EPPO, 2010; EPPO, 2014
ChinaPresentCABI/EPPO, 2010; EPPO, 2014
-HeilongjiangPresentCABI/EPPO, 2010; EPPO, 2014
-Nei MengguPresentLi et al., 1998; CABI/EPPO, 2010; EPPO, 2014
Georgia (Republic of)PresentCABI/EPPO, 2010; EPPO, 2014
IndiaPresentCABI/EPPO, 2010; EPPO, 2014
-Jammu and KashmirPresentCABI/EPPO, 2010; EPPO, 2014
-Tamil NaduPresentCABI/EPPO, 2010; EPPO, 2014
-Uttar PradeshPresentCABI/EPPO, 2010; EPPO, 2014
JapanPresentCABI/EPPO, 2010; EPPO, 2014
-HokkaidoRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-HonshuRestricted distributionCABI/EPPO, 2010; EPPO, 2014
Korea, DPRPresentCABI/EPPO, 2010; EPPO, 2014
Korea, Republic ofPresentCABI/EPPO, 2010; EPPO, 2014
NepalPresentCABI/EPPO, 2010; EPPO, 2014
PakistanRestricted distributionCABI/EPPO, 2010; EPPO, 2014
PhilippinesPresentCABI/EPPO, 2010; EPPO, 2014
Sri LankaPresentCABI/EPPO, 2010; EPPO, 2014

Africa

EthiopiaPresentCABI/EPPO, 2010
KenyaPresentCABI/EPPO, 2010; EPPO, 2014
MalawiPresentCABI/EPPO, 2010; EPPO, 2014
South AfricaRestricted distributionCABI/EPPO, 2010; EPPO, 2014
SwazilandPresentCABI/EPPO, 2010; EPPO, 2014
TanzaniaPresentCABI/EPPO, 2010; EPPO, 2014
UgandaPresentCABI/EPPO, 2010; EPPO, 2014
ZambiaPresentCABI/EPPO, 2010; EPPO, 2014
ZimbabwePresentCABI/EPPO, 2010; EPPO, 2014

North America

CanadaRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-British ColumbiaPresentCABI/EPPO, 2010; EPPO, 2014
-ManitobaPresentCABI/EPPO, 2010; EPPO, 2014
-Newfoundland and LabradorPresentCABI/EPPO, 2010; EPPO, 2014
-OntarioRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-QuebecPresentCABI/EPPO, 2010; EPPO, 2014
-SaskatchewanPresentCABI/EPPO, 2010; EPPO, 2014
USARestricted distributionCABI/EPPO, 2010; EPPO, 2014; EPPO, 2014
-AlaskaPresentCABI/EPPO, 2010
-CaliforniaPresentCABI/EPPO, 2010; EPPO, 2014
-FloridaPresentCABI/EPPO, 2010; EPPO, 2014
-HawaiiRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-IdahoPresentCABI/EPPO, 2010; EPPO, 2014
-IllinoisPresentCABI/EPPO, 2010; EPPO, 2014
-IowaPresentCABI/EPPO, 2010; EPPO, 2014
-MarylandPresentCABI/EPPO, 2010; EPPO, 2014
-MichiganPresentEPPO, 2014
-MinnesotaPresentCABI/EPPO, 2010; EPPO, 2014
-MontanaPresentTaylor and Schwandt, 1998; Taylor and Walla, 1999; CABI/EPPO, 2010; EPPO, 2014
-NebraskaPresentCABI/EPPO, 2010; EPPO, 2014
-North DakotaPresentBarnes et al., 2014
-OhioPresentCABI/EPPO, 2010; EPPO, 2014
-OklahomaPresentCABI/EPPO, 2010; EPPO, 2014
-OregonPresentCABI/EPPO, 2010; EPPO, 2014
-South DakotaPresentBarnes et al., 2014
-VermontPresent2000Pfister et al., 2000; CABI/EPPO, 2010; EPPO, 2014
-VirginiaPresentCABI/EPPO, 2010; EPPO, 2014
-WashingtonPresentCABI/EPPO, 2010; EPPO, 2014

Central America and Caribbean

Costa RicaPresentCABI/EPPO, 2010; EPPO, 2014
GuatemalaPresentCABI/EPPO, 2010; EPPO, 2014
HondurasPresentCABI/EPPO, 2010; EPPO, 2014
JamaicaPresentCABI/EPPO, 2010; EPPO, 2014
NicaraguaAbsent, unreliable recordCABI/EPPO, 2010; EPPO, 2014

South America

ArgentinaPresentCABI/EPPO, 2010; EPPO, 2014
BoliviaPresentCABI/EPPO, 2010
BrazilPresentCABI/EPPO, 2010; EPPO, 2014
-ParanaPresentCABI/EPPO, 2010; EPPO, 2014
-Sao PauloPresentCABI/EPPO, 2010; EPPO, 2014
ChileWidespreadCABI/EPPO, 2010; EPPO, 2014
ColombiaPresentCABI/EPPO, 2010; EPPO, 2014
EcuadorRestricted distributionCABI/EPPO, 2010; EPPO, 2014
UruguayWidespreadCABI/EPPO, 2010; EPPO, 2014

Europe

AustriaPresentCABI/EPPO, 2010; EPPO, 2014
BelgiumPresent, few occurrencesIPPC, 2008; CABI/EPPO, 2010; EPPO, 2014
BulgariaRestricted distribution****CABI/EPPO, 2010; EPPO, 2014
CroatiaPresentGlavas et al., 1997; CABI/EPPO, 2010; EPPO, 2014
Czech RepublicRestricted distributionCABI/EPPO, 2010; EPPO, 2014
DenmarkPresentCABI/EPPO, 2010; EPPO, 2014
EstoniaRestricted distributionCABI/EPPO, 2010; Drenkhan et al., 2014; EPPO, 2014
FinlandRestricted distributionCABI/EPPO, 2010; EPPO, 2014
FranceRestricted distributionLandmann, 2000; CABI/EPPO, 2010; EPPO, 2014; EPPO, 2014; Piou and Ioos, 2014
-France (mainland)Restricted distributionCABI/EPPO, 2010
GermanyPresent, few occurrences1983CABI/EPPO, 2010; EPPO, 2014; Heydeck et al., 2017
GreecePresentCABI/EPPO, 2010; Tsopelas et al., 2013; EPPO, 2014
-Greece (mainland)PresentCABI/EPPO, 2010
HungaryPresentCABI/EPPO, 2010; Barnes et al., 2011; EPPO, 2014
ItalyRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-Italy (mainland)Restricted distributionCABI/EPPO, 2010
LatviaPresentIPPC, 2009; EPPO, 2014Present: under eradication.
LithuaniaPresent, few occurrencesMarkovskaja and Treigiene, 2009; CABI/EPPO, 2010; EPPO, 2014; IPPC, 2016
NetherlandsPresentNPPO of the Netherlands, 2013; IPPC, 2008; CABI/EPPO, 2010; EPPO, 2014
NorwayPresentSolheim and Vuorinen, 2011; EPPO, 2014
PolandRestricted distribution1990Kowalski and Jankowiak, 1998; CABI/EPPO, 2010; EPPO, 2014; Boron et al., 2016
PortugalRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-AzoresPresentCABI/EPPO, 2010; EPPO, 2014
-Portugal (mainland)PresentCABI/EPPO, 2010
RomaniaRestricted distributionCABI/EPPO, 2010; EPPO, 2014
Russian FederationPresentBarnes et al., 2008; EPPO, 2014
-Southern RussiaPresentEPPO, 2014
SerbiaPresentCABI/EPPO, 2010; EPPO, 2014
SlovakiaPresentCABI/EPPO, 2010; EPPO, 2014
SloveniaPresentMacek, 1975; Jurc, 2007; CABI/EPPO, 2010; EPPO, 2014; EPPO, 2014
SpainRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-Spain (mainland)Restricted distributionCABI/EPPO, 2010
SwitzerlandPresent, few occurrences****CABI/EPPO, 2010; EPPO, 2014; Queloz et al., 2014
UKRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-England and WalesRestricted distribution1954CABI/EPPO, 2010; EPPO, 2014
-ScotlandPresent, few occurrencesCABI/EPPO, 2010; EPPO, 2014
UkrainePresentBarnes et al., 2008; CABI/EPPO, 2010; EPPO, 2014
Yugoslavia (Serbia and Montenegro)Present

Oceania

AustraliaRestricted distributionCABI/EPPO, 2010; EPPO, 2014
-New South WalesPresentCABI/EPPO, 2010; EPPO, 2014
-QueenslandPresentCABI/EPPO, 2010; EPPO, 2014
-TasmaniaPresentCABI/EPPO, 2010; EPPO, 2014
-VictoriaPresentCABI/EPPO, 2010; EPPO, 2014
New ZealandWidespreadCABI/EPPO, 2010; EPPO, 2014
Papua New GuineaPresentCABI/EPPO, 2010; EPPO, 2014

Risk of Introduction

Top of page The main phytosanitary risk is in the export/import of diseased plant material between countries. There are import restrictions on M. pini in many countries including Australia, New Zealand and the Czech Republic (Eldridge and Simpson, 1987; Jankovsky, 1998). Countries likely to be most affected are those with a heavy reliance on susceptible pines for commercial forestry such as Chile and New Zealand in which P. radiata comprises 81% and 90% of plantations, respectively (1999/2000 figures). It is not yet known if isolates of the pathogen from different countries differ in virulence. Until this is known it is imperative to restrict the transfer of isolates even into countries that already harbour the disease (Bradshaw et al., 2000).

Hosts/Species Affected

Top of page Dothistroma blight is mainly a disease of the genus Pinus. Pinus species vary in their susceptibility to M. pini infection. Pinus radiata is the most economically important and is highly susceptible. Some species, such as P. radiata, develop resistance with age and maturity whilst others such as P. ponderosa remain susceptible throughout their lifespan (Gibson, 1972).

Relative susceptibilities of some of the species listed are shown (Kershaw et al., 1988):
- Very highly susceptible: P. attenuata
- Highly susceptible throughout lifespan: P. jeffreyi, P. nigra subsp. laricio, P. ponderosa
- Highly susceptible but resistance increases with age: P. muricata, P. radiata
- Moderately susceptible: P. canariensis, P. lambertiana, P. pinaster
- Slightly susceptible: P. contorta, P. elliottii, P. hartwegii, P. monticola, P. nigra subsp. nigra
- Slightly susceptible and usually infected only when near diseased pines: Larix decidua, Picea sitchensis, Pseudotsuga menziesii
- Very slightly susceptible: Pinus ayacahuite, P. coulteri, P. michoacana, P. montezumae, P. patula, P. pseudostrobus, P. sabiniana, P. serotina, P. strobus, P. sylvestris, P. taeda, P. torreyana

For citations for host range of M. pini: see Dubin and Walper (1967); Cobb and Miller (1968); Basset (1969); Arthaud (1972); Skelly (1972); Shelbourne (1974); Ito et al. (1975); Wheeler et al. (1976); Sutton (1980); Lundquist and Roux (1984); Peterson (1984); Roux (1984); Lang (1987); Lang and Karadzic (1987); Kershaw et al. (1988); Fonseca Neves et al. (1990); Ivory (1994); Hunt (1995); Taylor and Walla (1999); Pfister et al. (2000); and Pehl and Wulf (2001).

Growth Stages

Top of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage

Symptoms

Top of page The most characteristic symptom of this disease is 1-3 mm wide brick-red bands that appear on the needles and persist after the needles have withered and turned brown. The red coloration is due to the presence of a mycotoxin, dothistromin. Generally the red zone is distinctly marked off from the rest of the needle and roughly spherical black fruiting bodies (stromata) erupt in the red infected band. Adjacent to the red band are bands of yellow necrotic needle tissue. Flanking these can sometimes be seen a band of more intense green pigmentation, representing increased lignification of the needle tissue as a defence response. The end of the needle dies beyond the point of infection and the whole needle may develop extensive necrosis (browning) 2-3 weeks after the first appearance of symptoms. Diseased needles drop prematurely (Edwards and Walker, 1978; Kershaw et al., 1988).

In a few reported cases the characteristic red pigment is not seen (Pehl and Butin, 1992; Ivory, 1994). On Pinus radiata, diseased needles may show various degrees of damage, from clear red bands around the needle to complete discoloration and death of the needle (Edwards and Walker, 1978). On Austrian pine (P. nigra) early symptoms include deep green bands and yellow and tan spots on needles. Later the spots and bands turn brown to reddish-brown (Peterson and Graham, 1974).

The first symptoms are found on needles of lower branches and the pathogen gradually moves up the crown. In some cases the disease starts in inner parts of lower branches and moves up the inner crown then subsequently outwards along the branches (Marks et al., 1989). Successive years of severe disease and premature defoliation result in decreased growth and, in extreme cases, death of the tree.

List of Symptoms/Signs

Top of page
SignLife StagesType
Leaves / abnormal colours
Leaves / abnormal leaf fall
Leaves / fungal growth
Leaves / necrotic areas

Biology and Ecology

Top of page Life Cycle and Reproductive Strategy

Conidia are passively transported in water droplets onto needles. Germ tubes form (one from each cell) and grow over the surface. There are conflicting reports about whether growth is directed towards stomata or whether it is random, although there is more tendency for directed growth in natural compared to artificial inoculations (Gadgil, 1967; Ivory, 1967; Peterson and Walla, 1978). An appressorium forms over a stomatal pore and a narrow infection peg grows into the plant. Septate hyphae branch out into the mesophyll and can be intra- and inter-cellular, but are restricted to necrotic tissues. Dead mesophyll cells adjacent to colonized areas suggest that host cells are killed in advance of the hyphae by a toxin (or by the host defence response). After 5-16 weeks (depending on environmental and host conditions) the host cells collapse and give rise to a visible lesion containing stromata (Ivory, 1972a; Peterson, 1973). In some parts of North America two growing seasons are required for the fungus to complete its life cycle (Taylor and Schwandt, 1998) but this does not seem to be the general rule.

Stromata mature and begin to produce conidia in spring. They are released in wet weather and can be dispersed by rain splash. The main period for infection is generally from late spring to late summer (May-August [Northern hemisphere] or November-February [Southern hemisphere]) (Gilmour, 1981) but the timing of spore dispersal can vary even within one country (Peterson and Harvey, 1976). Spore traps in Yugoslavia showed that conidia can be dispersed over a long period (7 months - spring to autumn) and particularly during periods of high humidity when conditions for infection are favourable (Karadzic, 1989). In spore trapping experiments in the USA, no conidia were collected on dry days, but even 0.23 cm of rain resulted in large numbers of spores being released within the crown of diseased trees (Peterson, 1973).

Ascospores are also produced, but for a shorter period of time, hence they are not considered to be such an important source of inoculum as conidiospores (Karadzic, 1989). On Pinus nigra in Germany only the conidial state was observed during the first year of infection, with ascostromata formed later, and it was proposed that the teleomorph is saprophytic whilst the anamorph is parasitic (Butin, 1985).

Factors Affecting Infection

In order to achieve infection, optimal conditions are required for fungal sporulation and growth. The major requirement is high humidity (Hunt, 1995). In the Northern hemisphere the amount of rainfall in June-September is a good indicator of the severity of disease (Peterson, 1973). Infections occur within the temperature range 5-26°C, with an optimum for conidial germination and stomatal entry of about 17°C.

Stromata formation, rather than germination and penetration, is inhibited in dry conditions. Hydrated conidia that land on a needle will germinate and penetrate regardless of the period of leaf wetness period that follows, provided that the temperature is suitable. However, the severity of disease depends on the length of the dry period following infection. The longer the dry period, the lower the disease severity and the longer taken for stromata to appear (Gadgil, 1977).

A 4-year study showed that a temperature of at least 10°C was required for infection, along with a period of high humidity/wetness for 15 hours. Infection only occurred at lower temperatures if the period of high humidity was extended (Gilmour and Crockett, 1972). In a later 3-year study there was no natural infection when the air temperature dropped below 7°C or when the leaf wetness was less than 10 hours (Gilmour, 1981).

Although a linear relationship has been found between light intensity and severity of disease, low light levels do not affect germination of conidia or early growth of the fungus on the needle surface. It is suggested that the response of the host to low light intensities, rather than that of the fungus, results in reduced levels of disease (Gadgil and Holden, 1976). Resistance is induced by shade treatment during the period 5-20 days after inoculation. On shaded foliage, penetration does not result in the development of disease symptoms (Ivory, 1972b).

Needle monoterpenes are more abundant in needles of younger trees than mature trees and stimulate spore germination and mycelial growth in vitro. However, the monoterpene composition profile does not vary with mature tree resistance (Franich et al., 1982). Conversely, a study of resistant needles from mature P. radiata showed that stomata are occluded with resinous material (surface wax). As well as providing a physical barrier to penetration of the fungus, the chemical nature of the occluding material is important as a pre-infectional chemical fungistasis factor. Oxidized resin acid derivatives from the surface wax of mature needles inhibited M. pini spore germination in vitro (Franich et al., 1983). In a similar study comparing surface wax from young pines of two different species, needles of susceptible P. nigra and resistant P. sylvestris contained about the same amount of wax (Walla and Peterson, 1976).

Highest disease levels on P. radiata in Australia occurred on poor soils (sulphur-deficient basalt), but were also influenced by other soil and topographic factors (Eldridge et al., 1981). Infections were also more severe in stands treated with high levels of nitrogen fertilizer which also led to a slower growth rate (Lambert, 1986).

Survival Strategies

Conidia remain viable for 2-6 months on diseased foliage lying on the forest floor under damp conditions (Gadgil, 1970). However, if the needles are stored dry, viability is retained for up to 11 months (Gibson et al., 1964).

Evidence for Adaptability

M. pini (or its anamorph) is believed to be endemic to America and Europe, and was not noticed in Africa or Australasia until extensive planting of exotic pine species had occurred. However, since the fungus arrived in New Zealand there has been no evidence of further adaptability. The genetic diversity is low and all isolates tested using molecular methods, including samples isolated in the 1960s and the 1990s, appear to be clonal (Hirst et al., 1999).

M. pini has recently been spreading in Europe (Pehl and Wulf, 2001). Differences in spore germination, growth rates and dothistromin production (in culture) have been found between isolates from different countries (Karadzic, 1987b; Bradshaw et al., 2000). Whether these isolates differ in virulence is not known.

Notes on Natural Enemies

Top of page Although there are no established examples of natural enemies, two studies are worthy of mention.

Pinus radiata trees with ectomycorrhizae of the Russulaceae were protected from attack by M. pini. Extracts from these fungi or from needles of mycorrhizal trees strongly inhibited spore germination of the pathogen, whilst needle extracts from non-mycorrhizal trees were not inhibitory (Garrido et al., 1982).

Ivory attempted to find natural enemies of M. pini. However, of 36 microorganisms isolated from P. radiata foliage only two fungi (Penicillium sp. and Periconia minutissim) and one bacterial species (unclassified) were antagonistic to M. pini in culture. The antagonism was not effective on pine needles (Ivory, 1972a).

Means of Movement and Dispersal

Top of page Airborne conidia are released and dispersed by rain splash for short distances. Spore trap experiments suggest that dispersal is limited to within trees or between closely placed trees - no spores were collected in traps placed 150 cm from the tree although abundant spores were collected within the tree canopy (Peterson, 1973).

Since the 1950s, the spread of M. pini has been rapid. From the first discovery of a severe outbreak in Tanzania in 1957, it was only 7 years before the disease was present in young Pinus radiata plantations in East and Central Africa. The spread of blight over long distances is not understood, but it is likely that wind, cloud and diseased materials (e.g. nursery stock) are possible transfer mechanisms (Gibson, 1974). Due to the geographic isolation of New Zealand it seems unlikely that wind/cloud dispersal of spores could account for its introduction there (first reported in 1964). The discovery that all New Zealand isolates tested are clonal suggests that a single introduction of the pathogen was responsible (Hirst et al., 1999). The disease is thought to have spread from New Zealand to Australia where it was first recorded in 1975. Strict quarantine regulations in Australia mean that introduction of diseased plant material is unlikely. Moist low-level airstreams flowing from New Zealand to Australia may have provided the necessary moisture and transport mechanism required for spore transport (Edwards and Walker, 1978; Marks et al., 1989). More recently both teleomorphic and anamorphic forms were found infecting an 8- to 10-year-old P. mugo stand in Germany in 1983. The probable cause of infection is the introduction of diseased P. nigra from a neighbouring country (Butin and Richter, 1983).

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Leaves Yes Yes Pest or symptoms usually visible to the naked eye
Seedlings/Micropropagated plants Yes Yes Pest or symptoms usually visible to the naked eye
Stems (above ground)/Shoots/Trunks/Branches Yes Yes Pest or symptoms usually visible to the naked eye
Plant parts not known to carry the pest in trade/transport
Bark
Bulbs/Tubers/Corms/Rhizomes
Flowers/Inflorescences/Cones/Calyx
Fruits (inc. pods)
Growing medium accompanying plants
Roots
True seeds (inc. grain)
Wood

Wood Packaging

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Wood Packaging not known to carry the pest in trade/transport
Loose wood packing material
Non-wood
Processed or treated wood
Solid wood packing material with bark
Solid wood packing material without bark

Impact

Top of page The economic impact of M. pini has been most serious in countries such as New Zealand, Chile and South Africa that have used Pinus radiata and other susceptible species as a major commercial forest crop. In New Zealand diseased foliage exceeds 10% on over 450,000 ha in the North Island (New and Griffith, 1989).

In P. radiata, the loss in wood volume growth is directly proportional to the average level of disease (estimated as percentage of crown affected) over a period of 8 years (Pas, 1981), i.e. 10% disease led to 10% loss of volume, 30% disease to 30% loss, etc. Other authors have published variations on these figures (Gibson, 1974). However, most agree that the impact is not considered significant until greater than 25% of the foliage becomes infected in 50% of the total number of trees in a stand (Kershaw et al., 1988) and at this stage fungicide spraying is considered economic.

Serious outbreaks have resulted in tree death in many parts of the world. There was 67% mortality in 7- and 8-year-old P. radiata in California, USA (Cobb et al., 1969) and Dothistroma needle blight caused complete failure of most P. ponderosa plantings in eastern states of the USA and up to 40% mortality of P. flexilis in Montana (Taylor and Schwandt, 1998). In Kenya in 1963, over 1500 hectares of P. radiata aged 1-5 years were so badly diseased that they were cut out and replanted with alternative species.

The annual cost of Dothistroma needle blight to the forestry industry in New Zealand was estimated to be NZ$6.1 m (about £2 m), in terms of direct control costs and residual growth loss (New and Griffith, 1989). In addition to these direct costs are the indirect effects. One report suggested that increased wood density occurs following severe defoliation, which would impact on processing costs (Harris and McConchie, 1978). Another indirect effect on wood yield is increased infection with secondary pathogens and pests, for example Sirex wasp (Sirex noctilio) which is more prevalent in stands with severe Dothistroma disease levels (Neumann et al., 1993).

Diagnosis

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The secretion of red-brown pigment (dothistromin) into agar by the cultured fungus is a useful diagnostic aid. A competitive ELISA test using monoclonal antibodies specific for dothistromin can be used for identification and quantification of the toxin (Jones et al., 1993; Bradshaw et al., 2000).

The DNA sequence of the variable internal transcribed spacer (ITS) region between the 18S and 5.8S ribosomal RNA genes was determined for a small collection of M. pini isolates from different countries. All isolates tested shared the same sequence except strains from Nebraska, Minnesota and Michigan, USA, that differed at 2 out of 144 nucleotide positions (Bradshaw et al., 2000); Genbank accession numbers AF462460, AF462459, AF013227. Using the same collection of isolates, microsatellite markers were developed that could be used to distinguish between geographically distinct isolates of M. pini (Ganley and Bradshaw, 2001).

A diagnostic protocol for Mycosphaerella pini is described in EPPO (2008).

Detection and Inspection

Top of page M. pini or its anamorph is detected on the basis of premature defoliation or needle browning of trees, followed by more detailed inspection of the needles for characteristic symptoms.

In New Zealand where the susceptible species Pinus radiata is grown commercially on a large scale, there is a well-established inspection system (Pas, 1981; Kershaw et al., 1988). Annual surveys are carried out of trees of susceptible age (2-15 years) in areas prone to pathogen attack. Ground surveys are carried out to determine the mean level of disease. Scoring is done visually by estimating the percentage of the crown that is affected and disease levels are estimated in 5% steps. If the overall level of stand infection is less than 15% then only a general estimation is necessary. If it is greater than 15% then an assessment of 100-200 trees is made, usually along a transect line. Aerial surveys are done by helicopter and allow rapid assessment of a whole forest. Rating is done in 5% steps but represents the average for the area rather than of individual trees. Because similar symptoms can be caused by other defoliating agents all stands rated at >15% are also checked in a ground survey. Because the accuracy of disease assessments depends on observer skill, forestry staff are given regular instruction and a training video is available (Information Officer, Forest Research Institute, Private Bag, Rotorua, New Zealand).

Similarities to Other Species/Conditions

Top of page Other Conditions

Symptoms may be confused with adverse environmental conditions such as boron and sulphur deficiencies. However, these are usually quite distinctive as tips of needles, whole needles or whole stands become uniformly affected. Where damage is caused by M. pini there is usually less uniformity of the symptoms. Sometimes only portions of the needles are killed, healthy needles are seen alongside diseased needles, and diseased trees may be adjacent to unaffected trees. In less severe pathogen infestations only the lower part of the tree is affected. Eventually the diseased needles develop dark fruiting bodies (Edwards and Walker, 1978; Hunt, 1995).

Other Species

Lecanosticta acicola [Mycosphaerella dearnessii] causes Lecanosticta needle blight (brown-spot needle blight or brown spot disease). This gives similar needle cast symptoms to M. pini on pine trees, with needles affected mainly on lower levels of trees and with similar dark stromatic fruiting bodies that erupt through the epidermis. Conidia are similar in shape and size, but whilst M. pini conidia are hyaline, conidia of L. acicola are greenish-brown. L. acicola produces no red-brown pigment (dothistromin) and necrosis is predominantly in spots rather than bands across the needle. An illustrated comparison of M. pini and L. acicola needle disease has been published (Pehl and Wulf, 2001).

Cercoseptoria pini-densiflorae (teleomorph Mycosphaerella gibsonii) causes Cercospora needle blight (brown needle disease). Symptoms usually develop in the centre of the lower crown then spread upwards and outwards. The first symptoms are light green bands occurring on young needles, spreading over the whole needle, turning yellow and eventually grey-brown. Exposed sympodial conidiogenous cells of the anamorph are an important feature that distinguishes C. pini-densiflorae from both M. pini and M. dearnessii, which have acervular or loculate conidiomata. Conidia are typically curved or lunate, 20-60 µm long and may have melanin granules in the spore walls that impart a stiffened or rigid appearance. Ascostromata are rarely grouped but spread along the needle (Evans, 1984; Crous et al., 1990).

Prevention and Control

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Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Cultural Control and Sanitary Methods

Dothistroma needle blight is commonly found in nurseries (Evans and Oleas, 1983; Dick and Vanner, 1986; Ivory, 1990) and movement of the disease around the world is probably due largely to diseased planting material (Gibson, 1974), hence rigid quarantine procedures are required in those nurseries currently free of disease (Wardlaw and Phillips, 1990). Pine nurseries should be as far as possible from pine forests, as diseased planting stock may disseminate inoculum into new areas (Hunt, 1995).

The type of tree planted has a significant effect on the level of disease. Some pine species are much more susceptible than others and some show resistance to infection upon maturity.

Because conidia remain viable for 6 months on damp leaf litter, Gadgil (1970) recommends waiting for this period after clear felling before replanting. This would not apply to species such as Pinus radiata that develop resistance at 15-20 years as they would not be expected to leave viable inoculum.

Standard commercial pruning removes many of the diseased branches and thereby decreases the inoculum in the forest environment (Kershaw et al., 1988). The beneficial effect of pruning on individual trees is disputed (Pas et al., 1984) although the effects can be influenced by their position in the stand. In a study of 5- to 7-year-old P. radiata, pruning reduced the level of infection within rows close to the edge of the plantation, possibly due to improved stand ventilation, but not those deeper within the plantation (Marks and Smith, 1987).

Host-Plant Resistance

Genetic variation in susceptibility to Dothistroma needle blight is documented in many species including P. ponderosa (Peterson, 1984), P. radiata (Wilcox, 1982), P. flexilis (Taylor and Schwandt, 1998) and P. muricata (Ades and Simpson, 1991), therefore the use of trees with increased resistance is recommended in high-risk areas. An extensive selection and breeding programme with P. radiata has yielded Dothistroma-resistant breeds that are expected to show a 15% decrease in mean stand infection and a 56% reduction in chemical spraying costs (Carson et al., 1991).

Some pine species, such as P. radiata and P. muricata, develop resistance with age and maturity (usually at 15 years). This is a feature of the whole tree in that resistance is seen even on new needles of mature trees (Ivory, 1972b). Other highly susceptible species remain susceptible throughout life (e.g., P. attenuata, P. nigra, P. ponderosa).

Chemical Control

The main method of control is spraying with copper oxychloride fungicides that kill the spores. Over large areas this is achieved using fixed wing aircraft fitted with micronair atomisers. In a trial with 13-year-old P. radiata sprayed three times when the mean crown disease levels reached 25% the final wood yield was estimated to be 30-40 m³/ha more than that from unsprayed control trees (Kershaw et al., 1988). In New Zealand one springtime spray gives 2-3 years protection, although severely affected areas are sprayed twice (Ray and Vanner, 1988). Details are given in a handbook published by the New Zealand Forest Research Institute (Kershaw et al., 1988). Copper fungicides have been used in many countries including Hungary (Koltay, 2001), Yugoslavia (Karadzic, 1987a), Chile (Rack, 1986) and the USA (Peterson, 1981). Large-scale spraying in East Africa was largely discontinued due to problems with difficult topography of the forests and a shortage of suitable airstrips and aircraft (Gibson, 1974).

Due to the high cost, some authors question whether there is a financial benefit to spraying (Pas et al., 1984), although it is considered that spraying should be continued to address concerns about the potential health hazards to forestry workers posed by dothistromin toxin (Elliot et al., 1989).

Other fungicides, such as benomyl are also effective in controlling blight but are not economic to use (Gibson, 1974; Karadzic, 1987a).

References

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Ades PK, Simpson JA, 1991. Variation in susceptibility to Dothistroma needle blight among provenances of Pinus radiata var. radiata. Silvae Genetica, 40(1):6-13

Arthaud J, 1972. Scirrhia pini (Ascomycetes, Dothideaceae) present in the southwest [of France]. Bulletin de la Societe Linneenne de Bordeaux, 2:123-124

Barnes I, Kirisits T, Akulov A, Chhetri DB, Wingfield BD, Bulgakov TS, Wingfield MJ, 2008. New host and country records of the Dothistroma needle blight pathogens from Europe and Asia. Forest Pathology, 38(3):178-195. http://www.blackwell-synergy.com/loi/efp

Barnes I, Kirisits T, Wingfield MJ, Wingfield BD, 2011. Needle blight of pine caused by two species of Dothistroma in Hungary. Forest Pathology, 41(5):361-369. http://onlinelibrary.wiley.com/journal/10.1111/(ISSN)1439-0329

Barnes I, Walla JA, Bergdahl A, Wingfield MJ, 2014. Four new host and three new state records of Dothistroma needle blight caused by Dothistroma pini in the United States. Plant Disease, 98(10):1443. http://apsjournals.apsnet.org/loi/pdis

Barr ME, 1996. Planistromellaceae, a new family in the Dothideales. Mycotaxon, 60:433-442; 29 ref

Basset C, 1969. Larix decidua a new host for Dothistroma pini. Plant Disease Reporter, 53:706

Boron P, Lenart-Boron A, Mullett M, Sieber T, 2016. The distribution of Dothistroma septosporum and its mating types in Poland. Forest Pathology, 46(5):489-496. http://onlinelibrary.wiley.com/journal/10.1111/(ISSN)1439-0329

Bradshaw RE, Ganley RJ, Jones WT, Dyer PS, 2000. High levels of dothistromin toxin produced by the forest pathogen Dothistroma pini. Mycological Research, 104(3):325-332; 29 ref

Butin H, 1985. Development of the teleomorph and anamorph of Scirrhia pini Funk & Parker on needles of Pinus nigra Arnold. Sydowia, 38:20-27

Butin H, Richter J, 1983. Dothistroma needle blight: a new pine disease in the German Federal Republic. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes, 35(9):129-131

CABI/EPPO, 1998. Distribution maps of quarantine pests for Europe (edited by Smith IM, Charles LMF). Wallingford, UK: CAB International, xviii + 768 pp

CABI/EPPO, 2010. Mycosphaerella pini. [Distribution map]. Distribution Maps of Plant Diseases, No.April. Wallingford, UK: CABI, Map 419 (Edition 5)

Carson SD, Dick AMP, West GG, 1991. Benefits of the Dothistroma-Resistant breed of radiata pine. In: Allen JC, Whyte AGD, eds. New Directions in Forestry: Costs and Benefits of Change. Australia and New Zealand Institutes of Forestry Conference, Christchurch, New Zealand: 251-262

Cobb FW, Miller DR, 1968. Hosts and geographic distribution of Scirrhia pini - the cause of red band needle blight in California. Journal of Forestry, 66:930-933

Cobb FW, Uhrenholst B, Krohn RF, 1969. Epidemiology of Dothistroma pini needle blight on Pinus radiata. Phytopathology, 59:1021-1022

Crous PW, Wingfield MJ, Swart WJ, 1990. Shoot and needle diseases of Pinus spp. in South Africa. South African Forestry Journal, 154:60-66

Dick M, Vanner AL, 1986. Nursery diseases. Forest Pathology in New Zealand, No.16

Doroguin G, 1911. Une maladie crytogamique du pin. Bulletin de la Societe Mycologique de France, 27:105-106

Drenkhan R, Adamson K, Jürimaa K, Hanso M, 2014. Dothistroma septosporum on firs (Abies spp.) in the northern Baltics. Forest Pathology, 44(3):250-254. http://onlinelibrary.wiley.com/journal/10.1111/(ISSN)1439-0329

Dubin HJ, Walper S, 1967. Dothistroma pini on Pseudotsuga menziesii. Plant Disease Reporter, 51:454

Edwards DW, Walker J, 1978. Dothistroma needle blight in Australia. Australian Forest Research, 8(2):125-137

Eldridge RH, Simpson JA, 1987. Development of contingency plans for use against exotic pests and diseases of trees and timber. 3. Histories of control measures against some introduced pests and diseases of forests and forest products in Australia. Australian Forestry, 50(1):24-36

Eldridge RH, Turner J, Lambert MJ, 1981. Dothistroma needle blight in New South Wales Pinus radiata plantation in relation to soil types. Australian Forestry, 44(1):42-45

Elliot GS, Mason RW, Ferry DG, Edwards IR, Griffith JA, 1989. Dothistromin risk assessment for forestry workers. Workshop on forest health in the South Pacific, Rotorua. Special issue, 19:2-3

EPPO, 2014. PQR database. Paris, France: European and Mediterranean Plant Protection Organization. http://www.eppo.int/DATABASES/pqr/pqr.htm

EPPO, 2018. Dothistroma pini does not occur in Germany. EPPO Reporting Service, 2018/181. Paris, France: EPPO. www.eppo.int

European and Mediterranean Plant Protection Organization, 2008. Mycosphaerella dearnessii and Mycosphaerella pini. Bulletin OEPP/EPPO Bulletin, 38(3):349-362. http://www.blackwell-synergy.com/loi/epp

Evans HC, 1984. The genus Mycosphaerella and its anamorphs Cercoseptoria, Dothistroma and Lecanosticta on pines. Mycological Paper, No.153:102 pp

Evans HC, Oleas AR, 1983. Pathology of Pinus radiata in Ecuador with special reference to Dothistroma. Tropical Pest Management, 29(4):316-320

Fonseca N, Laflamme G, 1997. Mycosphaerella pini (=Scirrhia pini), the perfect state of Dothistroma septospora: first observation in Portugal. Foliage, shoot and stem diseases. Proceedings of the IUFRO WP, 69-74

Fonseca-Neves N, Azevedo N de, 1990. Contribution to the knowledge on and control of diseases on Pinus pinea. Boletin de Sanidad Vegetal, Plagas, 16(2):447-453

Franich RA, Gadgil PD, Shain L, 1983. Fungistatic effects of Pinus radiata needle epicuticular fatty and resin acids on Dothistroma pini. Physiological Plant Pathology, 23(2):183-195

Franich RA, Gaskin RE, Wells LG, Zabkiewicz JA, 1982. Effect of Pinus radiata needle monoterpenes on spore germination and mycelial growth of Dothistroma pini in vitro in relation to mature tree resistance. Physiological Plant Pathology, 21(1):55-63

Funk A, 1979. Sexuality in Scirrhia pini. Bi-monthly Research Notes, 35(3):14

Funk A, Parker AK, 1966. Scirrhia pini n. sp., the perfect state of Dothistroma pini Hulbary. Canadian Journal of Botany, 44:1171-1176

Gadgil PD, 1967. Infection of Pinus radiata needles by Dothistroma pini. New Zealand Journal of Botany, 5:498-503

Gadgil PD, 1970. Survival of spores of Dothistroma pini. Report of Forest Research Institute, Rotorua, New Zealand, No. 48

Gadgil PD, 1977. Duration of leaf wetness periods and infection of Pinus radiata by Dothistroma pini. New Zealand Journal of Forestry Science, 7(1):83-90

Gadgil PD, Holden G, 1976. Effect of light intensity on infection of Pinus radiata by Dothistroma pini. New Zealand Journal of Forestry Science, 6(1):67-71

Ganley RJ, Bradshaw RE, 2001. Rapid identification of polymorphic microsatellite loci in a forest pathogen, Dothistroma pini, using anchored PCR. Mycological Research, 105(9):1075-1078; 13 ref

Garrido N, Becerra J, Marticorena C, Oehrens E, Silva M, Horak E, 1982. Antibiotic properties of ectomycorrhizae and saprophytic fungi growing on Pinus radiata D. Don I. Mycopathologia, 77(2):93-98

Gibson IAS, 1972. Dothistroma blight of Pinus radiata. Annual Review of Phytopathology, 10:51-72

Gibson IAS, 1974. Impact and control of dothistroma blight of pines. European Journal of Forest Pathology, 4:89-100

Gibson IAS, Christensen PS, Munga FM, 1964. First observations in Kenya on a foliage disease of Pines caused by Dothistroma pini Hulbary. Commonwealth Forestry Review, 43:31-48

Gilmour JW, 1981. The effect of season on infection of Pinus radiata by Dothistroma pini. European Journal of Forest Pathology, 11(5/6):265-269

Gilmour JW, Crockett F, 1972. Dothistroma pini project: monitoring of infection patterns in the field. Report of Forest Research Institute, Rotorua, New Zealand, No. 53

Glavas M, Diminic D, Hrasovec B, Margaletic J, 1997. Pests and diseases in Croatian forest nurseries recorded in 1996. Znanje za gozd. Zbornik ob 50. obletnici obstoja in delovanja Gozdarskega instituta Slovenije: Volume 1., 245-252; 12 ref

Goodwin SB, Dunkle LD, Zismann VL, 2001. Phylogenetic analysis of Cercospora and Mycosphaerella based on the internal transcribed spacer region of ribosomal DNA. Phytopathology, 91(7):648-658; 36 ref

Harris JM, McConchie DL, 1978. Wood properties of Pinus radiata infected with Dothistroma pini. New Zealand Journal of Forestry Science, 8(3):410-416

Heydeck P, Dahms C, Götz B, Hänisch A, Schumacher J, 2017. First record of Dothistroma needle blight (Dothistroma septosporum) in the northeast German lowlands. (Erster Nachweis der Dothistroma-Nadelbräune (Dothistroma septosporum) im Nordostdeutschen Tiefland.) Journal für Kulturpflanzen, 69(1):10-15. http://www.journal-kulturpflanzen.de

Hirst P, Richardson TE, Carson SD, Bradshaw RE, 1999. Dothistroma pini genetic diversity is low in New Zealand. New Zealand Journal of Forestry Science, 29(3):459-472; 20 ref

Hulbary RL, 1941. A needle blight of Austrian pines. Natural History Survey Bulletin, 21:231-236

Hunt RS, 1995. Common pine needle casts and blights in the Pacific Region. Forest Pest Leaflet - Pacific Forestry Centre, Canadian Forest Service, No. 43:7 pp.; 19 ref

IPPC, 2008. Scirrhia pini on Pinus sp. in public green, first detection in the Netherlands. IPPC Official Pest Report, No. NL-11/1. Rome, Italy: FAO. https://www.ippc.int/IPP/En/default.jsp

IPPC, 2008. Situation of Scirrhia pini in Belgium. IPPC Official Pest Report, No. BE-1/1. Rome, Italy: FAO. https://www.ippc.int/IPP/En/default.jsp

IPPC, 2009. First finding of the Scirrhia pini Funk & A. Parker in 2009. IPPC Official Pest Report, LVA-02/1. Rome, Italy: FAO. https://www.ippc.int/index.php?id=1110520&no_cache=1&type=pestreport&L=0

IPPC, 2016. Information on Pest Status in the Republic of Lithuania in 2015. IPPC Official Pest Report, No. LTU-01/2. Rome, Italy: FAO. https://www.ippc.int/

Ito K, Zinno Y, Suto Y, 1975. Dothistroma needle blight of pines in Japan. Bulletin of the Government Forest Experiment Station, Tokyo, No.272:123-140

Ivory MH, 1967. A new variety of Dothistroma pini in Kenya. Transactions of the British Mycological Society, 50:289-297

Ivory MH, 1972. Infection of Pinus radiata foliage by Scirrhia pini. Transactions of the British Mycological Society, 59(3):365-375

Ivory MH, 1972. Resistance to Dothistroma needle blight induced in Pinus radiata by maturity and shade. Transactions of the British Mycological Society, 59(2):205-212

Ivory MH, 1990. Needle diseases of pines in Nepal. Banko Janakari, 2(3):209-212

Ivory MH, 1994. Records of foliage pathogens of Pinus species in tropical countries. Plant Pathology, 43(3):511-518

Jankovsky L, 1998. Diseases of quarantine significance for tree species and plant protection in the Czech Republic. Lesnicka^acute~ Pra^acute~ce, 77(10):371-373

Jones WT, Harvey D, Jones SD, Fielder S, Debnam P, Reynolds PHS, 1993. Competitive ELISA employing monoclonal antibodies specific for dothistromin. Food & Agricultural Immunology, 5:187-197

Jurc D, 2007. Pines - Pinus spp. Diseases of needles. Lophodermium seditiosum, Mycosphaerella pini, Mycosphaerella dearnessii, Cyclaneusma minus. (Bori - Pinus spp. Bolezni iglic. Lophodermium seditiosum, Mycosphaerella pini, Mycosphaerella dearnessii, Cyclaneusma minus.) Gozdarski Vestnik, 65(7/8):321-336. http://www.dendro.bf.uni-lj.si/gozdv.html

Karadzic D, 1987. Effectiveness of some fungicides in the control of Dothistroma pini Hulbary in Pinus nigra plantations. Zastita Bilja, 38(1):15-31

Karadzic D, 1987. Effects of some ecological factors on the germination of spores and growth of mycelium of Scirrhia pini. Glasnik S^hacek~umarskog Fakulteta, Univerzitet u Beogradu, No. 69:93-118; 25 ref

Karadzic D, 1989. Scirrhia pini Funk et Parker. Life cycle of the fungus in plantations of Pinus nigra Arn. in Serbia. European Journal of Forest Pathology, 19(4):231-236

Karadzic DM, 1994. Picea omorika a new host of Dothistroma septospora. European Journal of Forest Pathology, 24(5):300-303

Kershaw DJ, Gadgil PD, Ray JW, Pas JBvd, Blair RG, 1988. Assessment and control of Dothistroma needle blight. FRI Bulletin, Forest Research Institute, New Zealand

Koltay A, 2001. Incidence of Dothistroma septospora (Dorog.) Morlet in the Austrian pine (Pinus nigra Arn.) stands in Hungary and results of chemical control trials. No^umlaut~ve^acute~nyve^acute~delem, 37(5):231-235; 11 ref

Kowalski T, Jankowiak R, 1998. First record of Dothistroma septospora (Dorog.) Morelet in Poland: a contribution to the symptomatology and epidemiology. Phytopathologia Polonica, No. 16:15-29; 25 ref

Lambert MJ, 1986. Sulphur and nitrogen nutrition and their interactive effects on Dothistroma infection in Pinus radiata. Canadian Journal of Forest Research, 16(5):1055-1062

Landmann G, 2000. Forest health in France: assessment for 1998 and new facts. Revue Forestiere Francaise, 52:9-22

Lang KJ, 1987. Dothistroma pini on young Norway spruce (Picea abies). European Journal of Forest Pathology, 17(4-5):316-317

Lang KJ, Karadzic D, 1987. Is Dothistroma pini a danger to Pinus sylvestris? Forstwissenschaftliches Centralblatt, 106(1):45-50

Li PF, Zhang XM, Hui EX, Liu ZF, Ge YX, Wang J, Wang DJ, Wu CY, 1998. Spatial distribution of needle blight disease of Pinus sylvestris var. mongolica. Journal of Forestry Research, 9:264-268

Lundquist JE, Roux C, 1984. Dothistroma needle blight of Pinus patula, P. radiata, and P. canariensis in South Africa. Plant Disease, 68(10):918

Macek J, 1975. Scirrhia pini, the pathogen of a new disease of Pine in Slovenia. Gozdarski Vestnik, 33:9-11

Markovskaja S, Treigiene A, 2009. New data on invasive pathogenic fungus Dothistroma septosporum in Lithuania. Botanica Lithuanica, 15(1):41-45

Marks GC, Smith IW, 1987. Effect of canopy closure and pruning on Dothistroma septospora needle blight of Pinus radiata D. Don. Australian Forest Research, 17(2):145-150

Marks GC, Smith IW, Cook IO, 1989. Spread of Dothistroma septospora in plantations of Pinus radiata in Victoria between 1979 and 1988. Australian Forestry, 52(1):10-19

Maschning E, Pehl L, 1994. Threat to native Pinus mugo by Dothistroma. AFZ, Allgemeine Forst Zeitschrift, 49(5):249-252

Morelet M, 1968. De Aliquibus in Mycologia Novitatibus (3 note). Bull. Soc. Sci. Nat. Archeol. Toulon. Var., 177:9

Neumann FG, Collett NG, Smith IW, 1993. The Sirex wasp and its biological control in plantations of radiata pine variably defoliated by Dothistroma septospora in north-eastern Victoria. Australian Forestry, 56(2):129-139

New D, Griffith JA, 1989. Forest health - an industry perspective of the risks to New Zealand's plantations. Special issue: Workshop on forest health in the South Pacific, Rotorua, 19:2-3

Pas JB van der, 1981. Reduced early growth rates of Pinus radiata caused by Dothistroma pini. New Zealand Journal of Forestry Science, 11(3):210-220

Pas JB van der, Bulman L, Horgan GP, 1984. Disease control by aerial spraying of Dothistroma pini in tended stands of Pinus radiata in New Zealand. New Zealand Journal of Forestry Science, 14(1):23-40

Pehl L, Butin H, 1992. Dothistroma septospora, a new fungus pest on Pinus mugo. AFZ, Allgemeine Forst Zeitschrift, 47(14):758-760; 15 ref

Pehl L, Wulf A, 2001. Mycosphaerella-needle fungi on pines - symptoms, biology and differential diagnosis. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes, 53(9):217-222; 4 ref

Peterson GW, 1973. Infection of Austrian and ponderosa pines by Dothistroma pini in Eastern Nebraska. Phytopathology, 63(8):1060-1063

Peterson GW, 1981. Control of Diplodia and Dothistroma blights of pines in the urban environment. Journal of Arboriculture, 7(1):1-5

Peterson GW, 1984. Resistance to Dothistroma pini within geographic seed sources of Pinus ponderosa. Phytopathology, 74(8):956-960

Peterson GW, Graham DA, 1974. Dothistroma needle blight of Pines. Forest Pest Leaflet, Forest Service, US Department of Agriculture, No. 143:5 pp

Peterson GW, Harvey GM, 1976. Dispersal of Scirrhia (Dothistroma) pini conidia and disease development in a shore pine plantation in Western Oregon. Plant Disease Reporter, 60(9):761-764

Peterson GW, Walla JA, 1978. Development of Dothistroma pini upon and within needles of Austrian and ponderosa pines in eastern Nebraska. Phytopathology, 68(10):1422-1430

Pfister SE, Halik S, Bergdahl DR, 2000. Dothistroma needle blight, caused by Dothistroma septospora, of Pinus spp. in Vermont. Plant Disease, 84(6):706; 1 ref

Piou D, Ioos R, 2014. First report of Dothistroma pini, a recent agent of the dothistroma needle blight, on Pinus radiata in France. Plant Disease, 98(6):841-842. http://apsjournals.apsnet.org/loi/pdis

Punithalingam E, Gibson IAS, 1973. Scirrhia pini. CMI Descriptions of Pathogenic Fungi and Bacteria, No. 368. Wallingford, UK: CAB International

Queloz V, Wey T, Holdenrieder O, 2014. First record of Dothistroma pini on Pinus nigra in Switzerland. Plant Disease, 98(12):1744. http://apsjournals.apsnet.org/loi/pdis

Rack K, 1986. On the seasonal release of Dothistroma pini conidia in Pinus radiata plantations in southern Chile. European Journal of Forest Pathology, 16(1):6-10

Ray JW, Vanner AL, 1988. Improvements in the technology of Dothistroma control. What's New in Forest Research, No. 169:4 pp

Roux C, 1984. The morphology of Dothistroma septospora on Pinus canariensis from South Africa. South African Journal of Botany, 3(6):397-401

Shelbourne CJA, 1974. Recent investigations of wood properties and growth performance in Pinus muricata. New Zealand Journal of Forestry, 19:13-45

Skelly JM, 1972. Dothistroma pini found infecting Scotch pine, Pinus sylvestris, a first report. Phytopathology, 62:671-672

Solheim H, Vuorinen M, 2011. First report of Mycosphaerella pini causing red band needle blight on Scots pine in Norway. Plant Disease, 95(7):875. http://apsjournals.apsnet.org/loi/pdis

Sutton BC, 1980. The Coelomycetes. Fungi imperfecti with pycnidia, acervuli and stromata. Wallingford, UK: CAB International

Taylor JE, Schwandt JW, 1998. Dothistroma needle blight of limber pine in Montana. Forest Health Protection Report - Northern Region, USDA Forest Service, No. 98-4:7 pp.; 13 ref

Taylor JE, Walla JA, 1999. First report of Dothistroma septospora on native limber and whitebark pine in Montana. Plant Disease, 83(6):590; 1 ref

Thyr DD, Shaw CG, 1964. Identity of the fungus causing redband disease on pines. Mycologia, 56:103-109

Tsopelas P, Barnes I, Soulioti N, Wingfield MJ, 2013. Dothistroma septosporum identified in Greece on Pinus brutia and Pinus nigra plantations. Plant Disease, 97(9):1247-1248. http://apsjournals.apsnet.org/loi/pdis

Walla JA, Peterson GW, 1976. Dothistroma pini and Diplodia pinea not affected by surface wax of pine needles. Plant Disease Reporter, 60(12):1042-1046

Wardlaw T, Phillips T, 1990. Nursery diseases and their management at the Forestry Commission nursery, Perth. Tasforests, 2(1):21-26; 2 ref

Wheeler NC, Kriebel HB, Lee CH, Read RA, Wright JW, 1976. 15-year performance of European Black Pine in provenance tests in north central United States. Silvae Genetica, 25:1-6

Wilcox MD, 1982. Genetic variation and inheritance of resistance to Dothistroma needle blight in Pinus radiata. New Zealand Journal of Forestry Science, 12(1):14-35

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